Hi

You can estimate the yield of your purification by two indices.

1- Recovery (Yield)= Total Units in purified fraction X 100/Total Units in crude extract.

2- Purification= Specific activity in each stage/Specific activity of crude extract.

To complete this you must perform a densitometry on your SDS GEL. You can use a free version of densitometry for instance:

http://www.gelanalyzer.com/download.html

However there are stronger softwares available if you're ready to pay for.

As Mahdi Hashempour Sadeghian said, you can perform densitometry of the Coomassie-stained gel in order to measure the amount of protein in the band that you assume to be your protein of interest. Measure the "volume" of the entire band and subtract the background from an area of the gel with the same level of background but no protein bands. Do this for each lane. Divide the background-corrected volume by the number of microliters of the original fraction that was loaded in the lane. Then multiply that figure by the total volume of the original fraction. This is the amount of protein in each fraction, expressed in arbitrary units of stain density. From this you can calculate the yield based on the stain density units in the crude extract.

If you have a pure sample of the protein, you can use it as a standard to relate stain density to amount of protein.

You can try to do this with a Western blot, but it is more difficult to be quantitative. You have to be careful about the linearity of signal to amount of protein.

You can do it by densitometry like Adam Shapiro suggested above - it is done commonly. You can either use software which comes with the imager that you took the image on, or general graphical software like ImageJ.

But the key requirement is that you need to relate the density of an observed band on the gel with the amount of protein you loaded onto the gel. So, if you still have your final pure sample - you can measure the protein concentration with techniques such as A280 spectrophotometry or Bradford reagent etc.

Hi Elia,

I agree with Adam B Shapiro , the linearity of signal to amount of protein will have to be accounted for.

Another technique is that, if you know the extinction coefficient of your pure protein, you can use a nanodrop to quantify the amount of protein.

I hope this helps.

use high/pure grade BSA known amount 100ng,250ng, 500ng,1ug, 2ug per well along with crude extract; 2-3 different dilution per lane. estimate based on BSA intensity (increasing amount) vs specific band in crude extract. you will have rough idea how much protein of interest is there in whole cell extract. you can also use UNScan IT for predicting density of BSA and you can extrapolate band of interest by using standard curve generated by BSA.

Hi Elia Scheja , your protein in chromatography fraction seems much clear compared to other fractions. You can easily quantify the purified protein using various techniques few of them are BCA(Bicinchoninic Acid Assay) which is a fast and reliable technique can be done using a 96 well plate and microplate reader. Also bradford assay which is again a fast and reliable technique can aslo be done in 96 well plate and as Anil Singh mentioned you need to have a standard usually BSA and few sets of concentration gradients (this conc. can be of your choice).

I am here attaching a link for protein qunatification using bradfor assay https://www.hivebench.com/protocols/9463 .

Best Wishes.

To add to above, it would be best to use your protein to check for staining linearity. Judging by your gel, you don't need to use as much of some of your samples as you did. Using your apple analogy, I would suggest 1 apple worth of sample 1; 3 apples worth of samples 2 and 3; then 2, 2.5, 3, 3.5 and 4 apples worth of purified protein sample. Using less of samples 2 and 3 will help resolve your protein of interest from the one that runs just below it. If you plan on using this method for estimating how much pure protein in the future, I also suggest that you write down your staining and destaining protocol in detail - composition, volume and time intervals of stain and of destain, how many times you change the destain, note down the setting that you use on the shaker and if the shaker is at room temperature or if you are sharing a shaker that has to be in a cold room for the work of other lab-mates. Then, you would not have to use so many lanes to establish linearity so carefully in future gels.

As others have commented, you can use the colour intensity in your stained gel as estimate of protein amount. However, because of the high concentration of protein in a gel band, it is highly questionable whether Lambert-Beer's law still applies. Thus, estimates may be severely biased.

Alternatively, you can measure the protein by any of the common methods, Bradford's assay and A280 are quick, with A280 you can even recover the sample.

So, measure the protein concentration (mg/mL) and the volume (mL) of your original sample and the recovered peak after purification and calculate the amount of protein (mg) in both. Then mgr/mgo * 100% is the recovery of total protein. This will always be smaller than 100%, after all, the purpose of purification is to get rid of extraneous proteins.

In addition, you need to measure the activity of your protein of interest (enzymatic activity, ELISA-signal, even band intensity if you have nothing better) and do the same calculation as above, to calculate recovery of your PoI. Usually, you will also loose some of your PoI in each purification step, in each step you have to balance recovery and purity. Sometimes, when you remove some inhibitor, the apparent yield of a step may, however, be larger than 100%.

Dividing the activity in a fraction by the amount of protein gives you the specific activity (say, kat/mg), which is a measure of purity.

When you purify a protein, all these measures should be tabulated so you can judge the efficiency of each purification step.